algae culture

Biotechnology Advances 30 (2012) 673–690

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Biotechnology Advances

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Research review paper

Microalgae biofuels: A critical review of issues, problems and the way forward

Man Kee Lam, Keat Teong Lee ⁎School of Chemical Engineering, Universiti Sains Malaysia, Engineering Campus, Seri Ampangan, 14300 Nibong Tebal, Pulau Pinang, Malaysia

⁎ Corresponding author. Tel.: +60 4 5996467; fax: +E-mail address: [email protected] (K.T. Lee).

0734-9750/$ – see front matter © 2011 Elsevier Inc. Alldoi:10.1016/j.biotechadv.2011.11.008

a b s t r a c t
a r t i c l e i n f o
Article history:Received 6 May 2011Received in revised form 16 November 2011Accepted 25 November 2011Available online 6 December 2011

Keywords:MicroalgaeEnergyBiofuelsBiomassSustainability

Culturing ofmicroalgae as an alternative feedstock for biofuel production has received a lot of attention in recentyears due to their fast growth rate and ability to accumulate high quantity of lipid and carbohydrate inside theircells for biodiesel and bioethanol production, respectively. In addition, this superior feedstock offers several en-vironmental benefits, such as effective landutilization, CO2 sequestration, self-purification if coupledwithwaste-water treatment and does not trigger food versus fuel feud. Despite having all these ‘theoretical’ advantages,review on problems and issues related to energy balance in microalgae biofuel are not clearly addressed untilnow. Base on thematurity of current technology, the true potential ofmicroalgae biofuel towards energy securityand its feasibility for commercialization are still questionable. Thus, this review is aimed to depict the practicalproblems that are facing the microalgae biofuel industry, covering upstream to downstream activities by acces-sing the latest research reports and critical data analysis. Apart from that, several interlink solutions to the prob-lems will be suggested with the purpose to bring current microalgae biofuel research into a new dimension andconsequently, to revolutionize the entire microalgae biofuel industry towards long-term sustainability.

© 2011 Elsevier Inc. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6732. Findings from LCA study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6743. Nutrient sources and culture methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6754. Raceway versus closed photobioreactor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6775. Harvesting of microalgae biomass . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6806. Drying of microalgae biomass . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6827. Microalgae lipid extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 683

7.1. Solvent extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6837.2. Supercritical fluid extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 684

8. Biodiesel production from microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6848.1. Homogeneous base and acid catalyst . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6858.2. Heterogeneous catalyst . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6858.3. In-situ transesterification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 685

9. Bio-oil production from microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68610. Bioethanol production from microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68611. CO2 bio-fixation using microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68712. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688Acknowledgment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688

1. Introduction

Since the last few decades, fossil fuels have become an integralpart of human daily lives. Specifically, fossil fuels are burned to

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produce energy for transportation and electricity generation, inwhich these two sectors have played a vital role in improvinghuman living standard and accelerating advance technological devel-opment. However, fossil fuels are non-renewable source that are lim-ited in supply and will one day be exhausted. In addition, burningfossil fuels have raised numerous environmental concerns, includinggreen house gas (GHG) effects which significantly contribute towards

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674 M.K. Lam, K.T. Lee / Biotechnology Advances 30 (2012) 673–690

global warming. Apart from that, as energy crisis is beginning to hitalmost every part of the world due to rapid industrialization andpopulation growth, the search for renewable energy sources has be-come the key challenge in this century in order to stimulate a moresustainable energy development for the future. Renewable energysources such as solar, wind, hydro, geothermal and energy from bio-mass and waste have been successfully developed and used bydifferent nations to limit the use of fossil fuels. Nevertheless, basedon a recent study from International Energy Agency (IEA), only ener-gy produced from combustible renewables and waste has the highestpotential among other renewable sources (International EnergyAgency, 2010). From the report, combustible renewables and wasteaccounted for 10.0% of the total primary energy supply, comparedto hydro energy 2.2% and other 0.7% (geothermal, solar, wind andheat). Hence, it was predicted that renewable energy from combusti-ble sources such as biodiesel will play a more crucial role as an alter-native renewable energy in the near future to further diversify theglobal energy sources.

Biodiesel is currently being recognized as a green and alternativerenewable diesel fuel that has attracted vast interest from re-searchers, governments, and local and international traders. Some ofthe advantages of using biodiesel instead of fossil diesel are that it isa non-toxic fuel, is biodegradable and has lower emission of GHGwhen burned in diesel engine (Demirbas, 2009). For the productionof first generation biodiesel, edible vegetable oils such as soybean,rapeseed, sunflower and palm oil are used as the main feedstock.However, the use of edible oils as energy source has raised a lot ofobjections from public and non-government organizations. Thus,second generation biodiesel derived from non-edible oils such asJatropha curcas L. appear as an attractive alternative feedstock forthe biodiesel industry. In fact, the use of jatropha oil in existing bio-diesel plant does not require major modification on the equipmentsand process flow, mainly because the oil has similar properties to ed-ible oils. However, jatropha oil does contain higher concentration offree fatty acid (FFA) that may require an additional pre-treatmentstep. Another advantage of using jatropha oil is that jatropha treescan grow easily on non-arable or wasteland. Nevertheless, regularirrigation, heavy fertilization and good management practices arerequired to ensure high oil yield (Lam et al., 2009). Due to theseweaknesses, the search for a more sustainable biodiesel feedstockcontinues and now focuses on microalgae.

Microalgae are recognized as one of the oldest living micro-organisms on Earth (Song et al., 2008). They grow at an exceptionalfast rate: 100 times faster than terrestrial plants and they can doubletheir biomass in less than one day (Tredici, 2010). Apart from that,some microalgae strains are able to accumulate large quantityof lipid inside their cells, in which the lipid can be converted to bio-diesel (Chisti, 2007). According to a recent study reported in theliterature, a realistic value of microalgae biomass production liesbetween 15 and 25 tonne/ha/year. With an assumption of 30% lipidcontent in microalgae cells (without optimizing the growth condi-tion), this is equivalent to a lipid production of 4.5–7.5 tonne/ha/year (Tsukahara and Sawayama, 2005). This amount is higher com-pared to the production of oil from soybean (0.4 tonne/ha/year),rapeseed (0.68 tonne/ha/year), oil palm (3.62 tonne/ha/year) andjatropha (4.14 tonne/ha/year) (Chisti, 2007; Lam and Lee, 2011). Inother words, culturing microalgae for biodiesel production requiresthe least land area and holds an important key feature for effectiveland utilization.

Apart from that, microalgae is also a superior feedstock forbioethanol production. Besides their high lipid content, some micro-algae also contain carbohydrates (generally not cellulose) that canbe used as carbon source or substrate for fermentation (Harun et al.,2010c). Even though research on the fermentation of microalgae bio-mass to bioethanol is still limited in the literature, but this processactually have a lot of advantages. In addition, microalgae are also

capable to fix CO2 from the atmosphere, flue gases or soluble carbon-ate into their cells during growth while simultaneously capturingsolar energy with efficiency 10–50 times greater than terrestrialplants; a golden opportunity for carbon credit program (Khan et al.,2009; Li et al., 2008b). Base on these evidences, microalgae have suc-cessfully positioned itself as one of the most promising biofuel feed-stock. Biofuels derived from microalgae are among third generationbiofuels that totally open up a new dimension in the renewable ener-gy industry. However it should be noted here that microalgae is notthe only feedstock that can be used for the production of third gener-ation biofuels. Other third generation biofuels are, for example bio-diesel produced from yeast and fungus (Nigam and Singh, 2011)and bioethanol produced via direct cellulose fermentation (consoli-dated bioprocessing) (Carere et al., 2008).

In recent years, the potential and prospect of microalgae for sus-tainable energy development have been extensively reviewed andmicroalgae are foreseen to be the fuel of the future. In fact, microalgaebiofuels have been placed globally as one of the leading researchfields which can bring enormous benefits to human beings and theenvironment. However, critical review on the problems facing theproduction of microalgae biofuels especially from thermodynamicperspective (mainly on energy balance) is not clearly addressed,resulting to unclear focus and direction on the development of micro-algae biofuels. Unlike other reviews reported in the literature, thispaper aims to access the actual scenario in microalgae biofuelsthrough the latest research reports rather than theoretical assess-ment. This review begins with the latest findings from life cycle as-sessment (LCA) to depict the actual problems and issues facing themicroalgae biofuels industry. Next, comprehensive comparisons be-tween terrestrial energy crops with microalgae will be evaluated,specifically in term of energy conversion efficiency. Apart from that,several technological modifications ranging from upstream (microal-gae culture, biomass harvesting and drying) to downstream processes(lipid extraction, biodiesel and bioethanol conversion techniques) aresuggested to provide a platform to drive more relevant and problemsolving research activities in this area of research for further improve-ment. It is hoped that this systematic and critical review will give aclear direction and guideline to researchers, policy makers, govern-ments, environmental protection agencies and business traders inpropelling the microalgae biofuels industry to become more viableand cost effective.

2. Findings from LCA study

Although microalgae biofuels (mainly biodiesel, bioethanol, bio-methane and bio-oil) have been foreseen to bring significant contri-bution in diversifying the global renewable energy sector, however,there is still a big question mark surrounding the sustainability ofthe industry for long term operation. Up to now, there is no commer-cial plant producing and processing microalgae biomass into biofuels.This has caused the lack of understanding in the overall process chainoperation. On the other hand, currently LCA is widely accepted as aneffective tool to guide and give a clear idea to researchers and policymakers on revealing the real potential of a particular product that isbeing evaluated. It can also be used to indicate if the production ofa particular product can lead to negative environmental phenomenasuch as eutrophication, global warming, ozone depletion, humanand marine toxicity, land competition, photochemical oxidation, etc.so that precautionary steps can be suggested to reduce the negativeimpacts. In addition, energy balance can be calculated to determineand justify the energy hotspot of all stages within the system bound-ary of the LCA.

Apparently, there are only a few LCA studies performed on micro-algae biofuels due to limited comprehensive data. Therefore, parame-ters related to microalgae biofuel production such as biomassproductivity, lipid content and downstream energy efficiency

675M.K. Lam, K.T. Lee / Biotechnology Advances 30 (2012) 673–690

(harvesting, drying and transesterification) are obtained based purelyon lab scale experimental data. Although the data used in thoseassessments might be irrelevant when applied to large-scale produc-tion, however, most of the studies have concluded that producingbiofuels from microalgae is an extremely energy intensive process.This finding is represented by the energy efficiency ratio (EER), de-fined as energy output to energy input is generally used to indicatethe sustainability energy index to produce a particular product, inwhich a ratio higher than 1 designates to net positive energy generat-ed and vice versa.

Table 1 shows a comparative study on EER for biofuels derivedfrom various feedstock including microalgae. However, it should benoted that the values presented in the table are a rough indicatorsince all the LCA studies were conducted based on different assump-tion and system boundary. From the table, it is rather unexpected tofind that biodiesel derived from oil bearing crops are much more en-ergy efficient compared to microalgae. All the EER value for biodieselderived from oil bearing crops are more than 1, whereas microalgaederived biodiesel have an EER value as low as 0.07. This quantitativeresults show that culturing of microalgae for biofuel productiondoes not necessary propel a positive output but worse still, couldpose a critical risk for unsustainable biofuels production. In addition,several issues such as reusability of water to re-culture microalgae,possibility of using contaminated wastewater as nutrients source,extraction and transesterification conversion efficiency, etc. havenot been clearly clarified in those LCA studies. If these factors aretaken into consideration, the EER value is expected to further diplower significantly. From the microalgae LCA studies, four of thekey energy intensive hotspots were identified: (1) nutrients source(Clarens et al., 2010), (2) photobioreactor design (Jorquera et al.,2010; Stephenson et al., 2010), (3) dewatering and biomass drying(Sander and Murthy, 2010) and (4) lipid extraction (Sander andMurthy, 2010; Stephenson et al., 2010). The following sections willdepict and explain these four key factors in detail.

Table 1Energy efficiency ratio (EER) for various energy crops and microalgae.

Feedstock EER Comment

Oil bearing cropsJatropha 1.92 With co-product productionJatropha 1.85 Without biogas productionJatropha 3.4 With biogas productionPalm oil 2.27 With co-product productionPalm oil 3.53 With co-product productionPalm oil 3.58 With co-product productionPalm oil 2.42 Without co-product productionRapeseed 1.44 With co-product productionRapeseed 5 Base on Chilean conditionsSunflower 3.5 Base on Chilean conditions

MicroalgaeChlorella vulgaris 0.35 Tubular photobioreactorChlorella vulgaris 1.46 RacewaysChlorella vulgaris 0.98 Sufficient nutrients condition and biomaChlorella vulgaris 3.54 Sufficient nutrients condition and biomaChlorella vulgaris 1.25 Low nitrogen culture and biomass are drChlorella vulgaris 4.34 Low nitrogen culture and biomass are noHaematococcus pluvaris 0.25–0.54 Haematococcus pluvaris strainNannochloropsis 0.09–0.12 Nannochloropsis strainNannochloropsis 1.08 Nannochloropsis strainNannochloropsis sp. 3.05 Raceways — the system boundary is limi

excluding dewatering, drying, extractionNannochloropsis sp. 1.65 Flat plat — the system boundary is limite

dewatering, drying, extraction and transeNannochloropsis sp. 0.07 Tubular photobioreactors — the system b

stage, excluding dewatering, drying, extrNot specified 3.33 Filter press as primary dewatering methoNot specified 1.77 Centrifuge as primary dewatering methoNot specified 1.06 Base case — inorganic (chemical) fertilizeNot specified 13.2 Conventional activated sludge as nutrien

3. Nutrient sources and culture methods

Culturing of microalgae at industrial scale for biofuels productionrequires substantial amount of nutrients, typically nitrogen (usuallyin the form of nitrate) and phosphorus (usually in the form ofortho-phosphate). These nutrients are normally from chemical or in-organic fertilizers that are used to achieve promising growth rate ofmicroalgae and to obtain bulk quantity of biomass. The use of chem-ical fertilizer has the advantage of reducing contamination in cultur-ing medium and thus promotes water reutilization to re-culturemicroalgae. However, a recent LCA study has pointed out that 50%of the overall energy use and GHG emission were associated with uti-lization of chemical fertilizers (Clarens et al., 2010). Chemical fertiliz-er production has been categorized as an energy intensive industry, inwhich 37 to 40 GJ of low heating value (LHV) natural gas will be con-sumed to produce 1 tonne of ammonia (inorganic nitrogen sources,N-fertilizer) (Rafiqul et al., 2005). Furthermore, 1.2 kg of carbon diox-ide (CO2) will be emitted for every 1 kg of ammonia produced (Kimand Dale, 2005). Hence, it is not surprising that the major source ofindustrial CO2 emission in the United States comes from ammoniamanufacturing plant (Kim and Dale, 2005). Thus, in a long run,using chemical fertilizers to culture microalgae for biofuel productionis definitely not sustainable.

Apart from that, culturing of microalgae is found to consume morechemical fertilizers than other oil-bearing crops as shown in Table 2.N-fertilizer is used in this assessment since N-fertilizer contributednearly 80–85% of the overall chemical fertilizers composition re-quired to culture microalgae. Based on the result presented in thetable, it is astonishing to find that culturing of microalgae requiredthe most fertilization when compared to other terrestrial oil-bearingplants. Oil palm plantation required the least fertilization, around83–87% lower than microalgae cultivation whereas sunflower,rapeseed and jatropha are 59–68%, 52–62% and 17–35% lower, re-spectively. This scenario implies that contributions of microalgae

Reference

Lam et al., 2009Achten et al., 2010Achten et al., 2010Lam et al., 2009Yee et al., 2009Pleanjai and Gheewala, 2009Pleanjai and Gheewala, 2009Yee et al., 2009Iriarte et al., 2010Iriarte et al., 2010

Stephenson et al., 2010Stephenson et al., 2010

ss are dried for extraction Lardon et al., 2009ss are not dried for extraction Lardon et al., 2009ied for extraction Lardon et al., 2009t dried for extraction Lardon et al., 2009

Razon and Tan, 2011Razon and Tan, 2011Batan et al., 2010

ted to cultivation stage,and transesterification stage.

Jorquera et al., 2010

d to cultivation stage, excludingsterification stage.


oundary is limited to cultivationaction and transesterification stage.


d (bioethanol is considered as second product) Sander and Murthy, 2010d (bioethanol is considered as second product) Sander and Murthy, 2010rs as nutrients source Clarens et al., 2010ts source Clarens et al., 2010

Table 2N-fertilizer consumption for various energy crops and microalgae.

Biodiesel feedstock N-fertilizer consumption(kg/kg oil)

Reference

Microalgae(Haematoccus pluvialis)

0.37a Razon and Tan, 2011

Microalgae(Nannochloropsis salina)

0.29b Batan et al., 2010

Jatropha 0.24c Achten et al., 2010Rapeseed 0.14d Iriarte et al., 2010Sunflower 0.12e Iriarte et al., 2010Palm 0.048f Pleanjai and Gheewala,

2009

a 0.032 kg N-fertilizer/kg oil for photobioreactor, 0.34 kg N-fertilizer/kg oil for race-way pond and 100% biodiesel conversion are assumed.

b 0.147 kg N-fertilizer/kg dry microalgae and 50% of lipid content are assumed.c 111 kg N-fertilizer/ha, 1695 kg seed/ha and 0.275 kg oil/kg seed are assumedd 68.2 kg N-fertilizer/tonne of seed and seed with 49% of oil content are assumed.e 57.1 kg N-fertilizer/tonne of seed and seed with 49% of oil content are assumed.f 7.79 kg N-fertilizer/tonne of fresh fruit bunch (FFB), 0.163 tonne oil/tonne FFB are

assumed.


biofuels toward environment conservation and energy security havereached a bottle neck as extremely high energy input is required dur-ing cultivation stage. Although the data provided in Table 2 is a roughindicator as the oil yield for each crop is highly dependent on localcondition and plant management practices, however, this findingshould not be taken lightly as these unforeseen circumstances maydrive the microalgae biofuels industry to a dead end. In addition, itshould be noted that the N-fertilizer consumption by microalgaewas calculated base on optimist condition, in which high lipid contentof 50 wt.% was assumed. If the lipid content is reduced to 22%(Boussiba et al., 1987; Su et al., 2010), N-fertilizer consumption willincrease to 0.67 kg/kg oil, an increment of 131% from the optimistscenario. Hence, recycling and reusing the excess nutrients in the cul-ture medium should be encouraged to improve the life cycle energybalance of microalgae biofuels. Perhaps, the main concern of thisapproach is the ability of microalgae to reuse the nutrients andgrow in a contaminated environment.

However, microalgae biomass yield (after lipid extraction) are su-perior compared to oil palm biomass, 45,500 kg/ha (Batan et al.,2010) for the former and 13,900 kg/ha for the latter (Shuit et al.,2009). Nevertheless, oil palm biomass (empty fruit bunches, fiber,shell, frond and trunk) were found to have high calorific value (18to 20 MJ/kg) which offer a good alternative and renewable sourcefor power generation (Lam et al., 2009; Shuit et al., 2009; Sumathiet al., 2008; Yee et al., 2009). Furthermore, oil palm biomass are cur-rently being explored for other uses, such as organic fertilizer, bio-plastic synthesis, pulp production, roughage source for ruminants,adsorbents for toxic gas and heavy metal removal (Shuit et al.,2009). In contrast, utilization of microalgae biomass for other applica-tions apart from biofuels production is still at the initial experimentalstage. Thus, it is a golden opportunity for researchers in this field toexplore other potential utilization of microalgae biomass and to fur-ther diversify more value added products that can generate revenuesfrom microalgae.

Due to the severe impact of chemical fertilizers towards the over-all energy balance in microalgae cultivation, there is an urgent needto search for alternative and low cost nutrient sources to ensurelong-term sustainability. In this case, using wastewater to culturemicroalgae appears as an attractive and economical alternative. Nor-mally, secondary and tertiary wastewaters contain significant amountof nitrate and ortho-phosphate which are not removed during prima-ry treatment. If these nutrients are to be removed, an additional 60 to80% of energy will be consumed in the wastewater treatment plant(Clarens et al., 2010; Maurer et al., 2003). Instead, these nutrientscan be used to culture microalgae and at the same time, microalgaeplay an important role as reagent to purify the wastewater. In

contrast, wastewater is not a suitable nutrient source for terrestrialcrops as the highly concentrated nutrients can easily leached outfrom soil and cause serious eutrophication of surface waters(Martijn and Redwood, 2005). For example, even trace amount of ni-trate in drinking water can be hazardous to human health and nitriteis exceptionally toxic to aquatic species (Martijn and Redwood,2005). Hence, culturing of microalgae in wastewater does not onlyoffers an inexpensive alternative to conventional way of wastewatertreatment, but also substantially reduced the need of chemical fertil-izers and their associated life cycle burden (Clarens et al., 2010).

From the environment and economic point of view, this is ulti-mately a win–win strategy in which waste stream from one processis used to generate alternative renewable bioenergies to mitigatethe current energy crisis. However, the real potential of using waste-water to culture microalgae is still uncertain and yet to be exploredsince wastewater is susceptible to bacteria and virus contamination.In the worst case scenario, these contaminations may devastate thewhole colony of microalgae and subsequently annihilate microalgaepopulation. Thus, frequent cleaning on raceway pond or photobior-eactor is mandatory to ensure optimum growth of microalgae. Inaddition, inconsistent nutrient composition in wastewater is also an-other uncontrolled factor which will directly retard the microalgaegrowth rate.

Although there are significant advantages of using wastewater toculture microalgae, nevertheless, research in this area is still very lim-ited. Up to now, only nearly 30% of published works on microalgaeculture are using wastewater as nutrients source whereas theremaining 70% use chemical fertilizers. This is because lab-scalemicroalgae culture limits the use of real industrial wastewater whilechemical fertilizer is easily available in the market and convenientlyprepared. Although promising results such as satisfactory growthrate and high lipid content can be achieved using chemical fertilizers,however, this method suffer serious limitation when up-scalingmicroalgae culture to industrial scale as discussed previously in theLCA study. Thus, the direction of research in this field has to be ur-gently diverted to ensure that sufficient data and information areprovided in the near future, typically related to ‘symbiosis’ and ‘rela-tionship’ between microalgae and wastewater.

Table 3 tabulated recent publications on different microalgae cul-turing methods for biodiesel production. From the table, Botryococcusbraunii, Chlorella protothecoides,and Chlorella vulgaris, have beenidentified as the most promising strains for biofuels production.Some of the interesting characteristics of these strains are: (1) canbe easily culture due to their simple unicellular cell, (2) high adapt-ability towards surrounding environment and (3) relatively highlipid content. Consequently, the survival rate of these strains is higherand thus, able to withstand other contaminants if cultured using in-dustrial wastewater. More importantly, these strains can be abun-dantly found in natural surroundings and therefore, related researchdata on these strains can be easily duplicated by researchers inevery part of this world for further improvement.

Studies have also shown that it is not necessary to culture micro-algae in medium with high concentration of nitrogen. By referringto Table 3, some reports highlighted that if microalgae are culturedunder nitrogen deficient condition, it can even lead to higher lipidaccumulation in microalgae cells, but at the risk of lower microalgaebiomass growth. This observation can possibly be explained as fol-lows: high nitrogen content in culture medium favors starch synthe-sis in microalgae cells to support their growth. However, whenmicroalgae are cultured in limited nitrogen medium, starch synthe-sis pathway is blocked and therefore photosynthetically fixedcarbon is redirected into fatty acid, leading to higher lipid accumu-lation in microalgae cells (Li et al., 2010). At the same time, starchwhich is the carbon and energy source in microalgae cells will getexhausted and hence, reduces photosynthetic efficiency. Therefore,microalgae growth will be retarded significantly and less biomass


are produced (Li et al., 2008a). In addition, there is possibility thatthe overall lipid productivity might even be lower in the case of ni-trogen deficient condition due to lower productivity of biomass(Huang et al., 2010). Consequently, bioethanol or biomethane pro-duction from microalgae biomass residue (after lipid extraction) issomehow limited since starch synthesis in microalgae cells areinhibited.

A recent LCA study has demonstrated that net negative energy bal-ance was attained in the process to produce biodiesel from dry micro-algae biomass when microalgae were cultured under nitrogen limitedmedium (Lardon et al., 2009). However, the study did suggest that ifthe lipid from microalgae were extracted through ‘wet extraction’method, it would greatly facilitate the overall process chain towardspositive energy balance. Nevertheless, extensive experimental worksare still required to justify this statement as only a handful of informa-tion about ‘wet extraction’ (e.g. type of solvent, effect of water contenttowards extraction efficiency and energy to evaporate the water afterextraction) is available in the literature at this moment. On the otherhand, other processing technologies such as pyrolysis, gasification orhydrothermal liquefaction seems to be a better way to utilize themicroalgae residue with low-carbohydrate content for subsequentbio-oil production. Through these technologies, the overall life cycleenergy balance of microalgae biofuels can be greatly improved asreported by Xu et al. (2011).

Generally, microalgae can be cultured using three methods:(1) phototrophic, (2) heterotrophic and (3) mixotrophic. Photo-trophic utilize light as energy source and CO2 as inorganic carbonsource whereas heterotrophic is independent of light and utilize or-ganic substrate (e.g. glucose, acetate, glycerol) as both energy andcarbon source (Mata et al., 2010). For mixotrophic culture, microalgaeare able to grow either via phototrophic or heterotrophic pathway,depending on the concentration of organic carbon sources and lightintensity (Mata et al., 2010). Up to now, only phototrophic methodis technically and economically feasible to culture microalgae in com-mercial scale, typically at outdoor environment where sunlight isabundant and free (Borowitzka, 1999). Apart from that, phototrophicmicroalgae are able to capture CO2 from flue gases and acts as a supe-rior carbon sink; an added advantage to the culture system. However,this method has its limitation especially in temperate countrieswhere suitable sunlight intensity is not always available throughoutthe year.

On the other hand, heterotrophic culture provides an immediatesolution to this problem as some of the microalgae strain can growunder dark environment. As shown in Table 3, relatively high lipidyield and biomass productivity can be attained through heterotrophicculture mode. In addition, the average lipid yield is around 26% higherthan phototrophic culture whereas the average biomass productivityis 265% higher. This phenomenon provides an absolutely good oppor-tunity for large scale production by using conventional fermentationbioreactor. Nevertheless, there are several key issues that need to beaddressed: (1) Not all microalgae strains can grow heterotrophically.Up to now, only C. protothecoides (Cheng et al., 2009; Xiong et al.,2008; Xu et al., 2006), C. vulgaris (Liang et al., 2009), Crypthecodiniumcohnii (Couto et al., 2010) and Schizochytrium limacinum (Johnsonand Wen, 2009) have been identified to be able to grow in total dark-ness and able to accumulate high quantity of lipid. (2) Serious con-tamination by other microorganism due to the presence of organicsubstrate (Chen et al., 2010). The contamination level is expected tobecome even more critical if wastewater is used as nutrients source.Although sterilization may provide a solution to this problem, howev-er, high energy consumption is required and further burdens theoverall process. (3) Increase in energy consumption and costs by add-ing organic substrate. At present, glucose is the most suitable organicsubstrate for growing microalgae heterotrophically due to its high en-ergy content, ~2.8 kJ/mol compared to acetate ~0.8 kJ/mol (Boyle andMorgan, 2009; Perez-Garcia et al., 2011). Nevertheless, glucose which

is obtained from sugar-based plant is equally important for humanconsumption and consequently may triggers food versus fuel feud.On the other hand, the potential of using glucose derived from ligno-celluloses biomass and glycerol which is the by-products from biodie-sel production are yet to be explored as an alternative and low-costcarbon substrate. (4) CO2 is released through microalgae respiration.This is an unfortunate scenario as heterotrophic microalgae does notfulfill the mandate of solar energy capture and unable to synergizeCO2 mitigation from atmosphere (Li et al., 2008a). Hence, more com-prehensive LCA studies and proactive research for heterotrophicmicroalgae culture is heavily required.

4. Raceway versus closed photobioreactor

Apart from nutrients supplement, microalgae cultivation systemalso plays an important role to determine the successfulness of the in-dustry. An effective culture system should consist of the following cri-teria: (1) effective illumination area, (2) optimal gas–liquid transfer,(3) easy to operate, (4) low contamination level, (5) low capital andproduction cost and (6) minimal land area requirement (Xu et al.,2009). Apparently, only raceway pond is implemented in commercialscale for microalgae cultivation to produce high-value nutritiousproducts for animal feed. The system consists of a paddle wheel toavoid microalgae biomass sedimentation and CO2 is sparged at thebottom of the raceway as carbon source (Stephenson et al., 2010).Although this system is relatively easy to operate and consume lessenergy, however high contamination level by undesired micro-organisms can eventually jeopardize the survival of microalgae.Apart from that, the system also experience high water loss due toevaporation (Schenk et al., 2008). Therefore, closed photobioreactoris introduced to overcome the limitations encountered in racewaypond. There are several designs of closed photobioreactor, such as air-lift tubular, flat plate, vertical-column, etc. The main advantage ofgrowing microalgae in a closed photobioreactor is it permits singlestrain culture, in which optimum growth condition is always main-tained to give high consistency in biomass and lipid productivity.Thus, closed photobioreactor has always attracted great interestfrom researches to further improve the operating conditions forimplementation in commercial scale.

Currently, a few LCA studies have been performed to evaluate en-ergy consumption in operating raceway pond and air-lift tubularphotobioreactor for microalgae cultivation as shown in Table 4. Nev-ertheless, the result is rather unexpected as raceway pond emergedas a more sustainable and economic way to culture microalgae eventhough optimum culture conditions (microalgae with high lipid pro-ductivity) are applied in air-lift tubular photobioreactor (Jorqueraet al., 2010; Stephenson et al., 2010). From the table, average energyinput to operate air-lift photobioreactor is around 350% higher com-pared to raceway pond. Despite the advantages of lower level of con-tamination and optimal use of cultivation area, air-lift tubularphotobioreactor consumed significantly high amount of electricityfor powering of pump so that sufficient mixing and optimum gas–liquid transfer are achieved. Base on currently available technology,air-lift photobioreactor is not up to commercialization stage unlessproper modifications are performed to reduce the overall operatingenergy consumption.

One of the plausible improvements is by designing an oscillatoryflow reactor rather than a tubular type (Stephenson et al., 2010). Os-cillatory flow reactor consists of equally spaced orifice plate baffles inwhich the baffles behave like stirred tanks that can give excellentmixing effect by creating vortices between orifice baffles and super-imposed oscillating fluid as shown in Fig. 1 (Harvey et al., 2003;Zheng et al., 2007). This type of reactor was previously tested for con-tinuous biodiesel production in which high biodiesel yield wasattained within a short reaction time due to the superior mixingintensity (Harvey et al., 2003). Thus, if the reactor is modified to

Table 3Different culture methods for various microalgae strains.

Strain Nutrients Culture condition Specific growthrate (day−1)

Biomassproductivity(g/L/day)

Biomassyield (g/L)

Lipidproductivity(g/L/day)

Total lipid extracted(wt.% of biomass)

Comment Reference

Botryococcusbraunii

Secondary treatedsewage

Air flowrate of 0.5 v/v/min 0.13 0.288 – – 17.85 High potential of using secondarytreated sewage from domesticwastewater to grow microalgae

Órpez et al., 2009Air flowrate of 1 v/v/min 0.14 0.346 – – 17.85

Botryococcusbraunii

Secondary domesticwastewater

100% of wastewater mediumwas used without dilution

0.11 0.034 0.48 – 36.14 Microalgae biodiesel productioncoupled with waste water treatmentappears as a good opportunity tocommercialize the process

Sydney et al., 2011

Botryococcusbraunii

Modified Chu 13 With supplement of 2%(v/v) CO2

– 0.043 0.9 – 22 The organism exhibited widerange of pH adaptability

Dayananda et al.,2007

Chlamydomonasreinhardtii

Wastewater Microalgae was culturedin biocoil

0.564 2 – 0.505 25.25 High lipid content and biomassproductivity can be attained throughbiocoil due to greater light exposureand intensity inside the polyvinyl tubing.

Kong et al., 2010

Chlorellaprotothecoides

Inorganic basalmedium

Heterotrophic culture withcorn powder hydrolysate ascarbon source

– 2.02 15.5 – 55.2 Higher lipid content was observedif microalgae cultured in heterotrophiccondition

Xu et al., 2006


Basal Medium Heterotrophic growth withglucose (24 g/L) as carbonsource and yeast (4 g/L) asfermentation media

– 7.3 51.2 – 50.3 High lipid was obtained throughheterotrophic growth

Xiong et al., 2008


Basal medium Heterotrophic culture withJerusalem artichoke (30 g/L)as carbon source

– 4.1 17 1.7 43 Jerusalem artichoke appears as a lowcost carbon source for heterotrophicmicroalgae culture

Cheng et al., 2009

Chlorella sp. Anaerobic digesteddairy manure

25× diluted digested dairymanure

0.409 – – – 13.7 Anaerobic digested manure served asa low cost nutrients source to culturemicroalgae while at the same time,microalgae provide a valuablesolution to refractory dairy waste.

Wang et al., 2010

Chlorella sp. Walne medium withurea as nitrogensource

Urea concentration of 0.025 g/Land microalgae was cultured inbatch culture mode for 6 days

0.86 – 0.464 0.051 66.1 Urea is relatively low cost compareto other inorganic nitrogen sources.High lipid content can be achievedthough urea-limited condition.

Hsieh and Wu,2009

Urea concentration of 0.20 g/Land microalgae was cultured inbatch culture mode for 6 days

1.42 – 2.027 0.11 32.6

Chlorella vulgaris Modified Fitzgeraldmedium

Normal nutrients condition(20 days)

– 0.043 0.86 0.0128 29.5 Two stage cultivation (N-sufficeintfollowed by N-limited condition)increased lipid content and graduallychanged the lipid composition fromfree fatty acid to mostly triacylceride(TG).

Widjaja et al.,2009

20 days of normal nutrientscondition followed by 17 daysof nitrogen limited condition

– – – – 44

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Chlorellavulgaris

Artificial wastewatermedium

Semi-continuous culture modewith daily replacement of freshmedium (until second phase)

– – 0.69 0.147 42 Microalgae biodiesel is comparativewith petroleum due to high potentialcredit to wastewater treatment

Feng et al., 2011

Chlorellavulgaris

Not specified(inorganic chemicals)

Heterotrophic culture with1% w/v glucose as carbon source

– 0.151 1.2 0.035 23 Glycerol, a by-product from biodieselindustry become a low cost carbonsource to culture microalgae

Liang et al., 2009

Heterotrophic culture with 1% w/vglycerol as carbon source

– 0.102 0.722 0.022 22

Heterotrophic culture with 2% w/vglycerol as carbon source

– 0.091 0.656 0.031 34

Chlorella vulgaris Guillard f/2 one Temperature as a controlling factortowards lipid yield:(a) 25 °C(b) 30 °C

0.140.14

–

–

–

–

0.020.008

14.715.9

Temperature play a significant rolein promoting lipid productivity

Converti et al.,2009

Nitrate as a controlling factortowards lipid yield:(a) 1.5 g/L NaNO3

(b) 0.375 g/L NaNO3

0.140.13

–

–

–

–

0.0080.02

5.915.3

Higher lipid yield can be achievedthrough nitrogen-limited condition

Chlorella vulgaris Artificial wastewater Semi-continuous cultivation(until second phase)

– – 0.69 0.147 42 Wastewater as nutrient sourceenhances cost effectiveness toproduce biodiesel.

Feng et al., 2011

Chlorella vulgaris Watanbe medium Nitrate sufficient condition 0.99 0.029 0.41 – 18 High lipid content can be achievedthrough heterotrophic culture

Illman et al., 2000Low nitrate condition 0.77 0.037 0.52 – 40

Dunaliella tertiolecta Erdschreiber Fluorescent light with lightintensity of 350 μE/m2.sand 4% CO2

– – 0.39 – 23.4 High tolerance towards CO2 level Tang et al., 2010

Isochrysis galbanaCCMP 1324

Modified Walne'smedium

Urea – 0.17 1.06±0.12 – 22.34±2.56 Urea as cheap nutrient source toculture microalgae

Su et al., 2007(NH4)2SO4 – 0.12 0.72±0.08 – 19.58±3.69NH4Cl – 0.12 0.72±0.056 – 14.32±3.33

Nannochloropsisoculata

Bold's Basal Medium Temperature as a controllingfactor towards lipid yield:(a) 20 °C(b) 25 °C

0.130.07

–

–

–

–

0.010.01

7.913.89

Temperature play a significant role inpromoting lipid productivity

Converti et al.,2009

Nitrate as a controllingfactor towards lipid yield(a) 0.3 g/L NaNO3

(b) 0.075 g/L NaNO3

0.130.1

–

–

–

–

0.010.016

7.8815.86

Higher lipid yield can be achievedthrough nitrogen-limited condition

Neochlorisoleabundans

Bristol medium Nitrogen-sufficient medium(30 °C with CO2 enrichment)

– 0.15 0.95 0.0378 28 High lipid content can be achievedthrough nitrogen-starvationcondition, however lowbiomass productivity was observed.

Gouveia et al.,2009

Nitrogen-limited medium(30 °C with CO2 enrichment)

– 0.03 0.15 0.0144 52

Neochlorisoleoabundans

Modified soilextract medium

Sodium nitrate concentrationof 3 mM

– 0.31 1.85 0.125 40 High lipid content can be achievedthrough nitrogen-limited condition

Li et al., 2008a

Sodium nitrate concentrationof 10 mM

– 0.63 3.15 0.098 15

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Table 4Energy consumption in different microalgae culture system.

Culture system Energy consumptiona

(GJ/tonne of biodiesel produced)Reference

Raceway 4–11 Lardon et al., 2009Raceway 13–15 Jorquera et al., 2010Raceway 22–30 Stephenson et al., 2010Raceway 53–158 Campbell et al., 2011Air-lift tubular 195–231 Stephenson et al., 2010Air-lift tubular 537 Jorquera et al., 2010

a Energy associated with electricity consumption to operate culture system:

Raceway — Paddle wheel and gas spargingFlat-plat — Pump and gas spargingAir-lift tubular — Air-lift pump and gas sparging.


become a photobioreactor, then energy consumption can be reducedbecause only minimal culture velocity is required even to achieveintense mixing effect. In addition, mass transfer of CO2 to culture me-dium can be further improved and enhances CO2 utilization by micro-algae. On the other hand, baffled system can be easily implemented incurrent raceway pond without major modification. A conceptual de-sign of the baffled raceway pond is shown in Fig. 2. From the figure,vortexes are created due to the baffled system and hence, enhancemixing intensity in the raceway pond. However, the performance ofthe baffled system in photobioreactor or raceway pond has yet to beevaluated, especially in term of fluid dynamic, CO2 mass transferefficiency, effect of turbulence towards microalgae growth, energyconsumption, and scalability.

Apart from the need to reduce substantial energy consumption inraceway or photobioreactor through innovative design, culturing ofmicroalgae is still heavily dependent on fossil fuels. This can be over-come by integrating microalgae farming with current energy cropsplantation to achieve a truly sustainable way of producing microalgae

Oscillatory flow

Baffle

Vortex

Fig. 1. Conceptual oscillatory flow reactor (OFR) for microalgae culture.Modified from Harvey et al. (2003).

biofuels. Energy crops, specifically referring to rapeseed, soybean, oilpalm, jatropha and sugar cane are currently used for food and fuelfeedstock. At the same time, the crops also produce huge amount ofbiomass which can subsequently be processed to a variety of prod-ucts, such as organic fertilizer, furniture, animal feed, etc. Apartfrom that, the biomass can also generate substantial amount of ener-gy if burned as fuel. The potential amount of energy generated aredepicted in Table 5; however, only biomass from oil bearing cropsare considered in order to ensure fair comparison (on the samebasis) with biodiesel produced from microalgae. From the table, it isworth to note that the energy generated from biomass can eitherfully or partially replace the energy required in operating racewayor tubular photobioreactor. Hence, the dependency on fossil fuel canbe reduced and this strategy appears to be a greener approach to cul-tivate microalgae for biofuels production.

In addition, some of the biomass contains high concentration oflignocellulose which can be converted to bioethanol or biomethanethrough a series of pre-treatment, hydrolysis, fermentative and an-aerobic digestion process. This approach is much safer and environ-mental friendly instead of burning the biomass directly as fuelwhich may cause serious air pollution. Apart from the biomass fromenergy crops, the retained microalgae biomass after lipid extractionalso has a huge potential for bioethanol or biomethane production.Recent studies have depicted that some microalgae strains containhigh concentration of carbohydrate (generally not cellulose) and pro-teins that can be used as carbon source or substrate for fermentation(Harun et al., 2010c). Base on this evidence, microalgae cultivationwill become more realistic with diversified products from microalgaebiomass or incorporation with current energy crops. Consequently,high energy input to operate photobioreactor can be offset, allowingmicroalgae farm to be self-sufficient in term of power requirement.In fact, there are other potential green technologies that can help tominimize the dependency of fossil fuel in microalgae farm, such assolar panel and wind turbines. Integration of these renewable ener-gies has yet to be discovered to revitalize a truly sustainable biofuelproduction from microalgae.

5. Harvesting of microalgae biomass

When microalgae culture has reached the stationary phase, thenext step is to separate the microalgae from water and to recovertheir biomass for downstream processing. Nevertheless, microalgaeharvesting process posed a challenging task to engineer since micro-algae are small size microorganism (generally, 1–20 μm) and sus-pended in liquid. Currently, there are several methods to harvestmicroalgae: (1) bulk harvesting – to separate microalgae from sus-pension, such as natural gravity sedimentation, flocculation and floa-tation and (2) thickening – to concentrate the microalgae slurry afterbulk harvesting, such as centrifugation and filtration (Chen et al.,2010). A recent LCA study has underlined that harvesting and dryingof microalgae biomass contributed significant energy input in produc-ing microalgae biodiesel (Sander and Murthy, 2010). Two types ofthickening methods (without prior bulk harvesting) were includedin the study, filter press and centrifugation; each of the method con-tributed 88.6% (equivalent to 122 GJ/tonne biodiesel) and 92.7%(equivalent to 239 GJ/tonne biodiesel), respectively to the entire en-ergy input of the LCA. The energy consumed is even comparable tothe energy used in operating tubular photobioreactor as indicated inTable 4. In this regard, the results show that bulk harvesting technol-ogies may play an important role in reducing the energy consumptionduring the thickening process of microalgae slurry. In short, the ener-gy consumed in harvesting and drying of microalgae biomass shouldnot be ignored as it may bring significant adverse effect towards theoverall energy balance in producing microalgae biofuels.

Up to now, centrifugation and filtration are still not energy-feasible methods to harvest microalgae in commercial scale. As

Fig. 2. Conceptual baffled system in raceway pond to culture microalgae.Modified from Chisti (2007).


aforementioned in the LCA study, centrifugation and filtrationmethods involve extensive energy consumption, and high capitaland maintenance cost resulting to unsustainable practice for longterm operation. On the other hand, flocculation offers a relativelylow energy way to harvest microalgae. This is because microalgaecells always carry negative charge which causes them to repel eachother and to be suspended in liquid for a long time although mixingis not provided. By introducing coagulant that is positively chargedinto the culture medium, the negative charge surrounding the micro-algae cells will be neutralized. At the same time, flocculant can beadded to promote agglomeration by creating bridges between theneutralized cells to become dense flocs and settle due to natural grav-ity (de Godos et al., 2011). Nevertheless, it is worth to mention thatflocculation with conventional multivalent salts (e.g. polyaluminiumchloride) will contaminate the microalgae biomass and consequently,may bring adverse effect towards the final product quality (Lee et al.,2010a). Although no scientific work or assessment is carried out tojustify this claim, however, flocculant toxicity should not be ignoredespecially if health related products are to be extracted out frommicroalgae biomass before being used for subsequent biofuelproduction.

In addition, conventional flocculation method also posed severaldisadvantages: (1) high dosage of multivalent salt is required toachieve satisfactory result, (2) produces large quantity of sludgethat increases the difficulty to dehydrate the biomass and (3) floccu-lation efficiency is highly dependent on pH level (Chen et al., 2010;Renault et al., 2009). Therefore, organic polymeric flocculant whichis biodegradable and less toxic offers an alternative and environmen-tal friendly way to aggregate suspended particles. Currently, theavailable organic polymeric flocculant in the market are mainlysynthetic, while flocculant derived from natural products such asstarches, polysaccharides and cellulose has attracted increasing inter-est from researchers (Renault et al., 2009). Nevertheless, polymericflocculant (especially cationic type) is ineffective for marine microal-gae due to inhibition by high ionic strength of seawater (Bilanovic

Table 5Average energy produced from different source of biomass (oil bearing crops).

Energycrops

Biomass Average energy produced(GJ/tonne biodiesel produced)

Reference

Rapeseed Straw 88.2 Janulis, 2004Soybean Soybean

meal74.2 Panichelli et al.,

2009Oil palm Empty fruit

bunches andfiber

95 Lam et al., 2009; Shuitet al., 2009

Jatropha Seed cake 103 Pleanjai and Gheewala,2009

et al., 1988). Hence, significant breakthrough findings for developingpolymeric flocculant is urgently required through intensive researchworks to further strengthen its potential usage in microalgae harvest-ing process.

On the other hand, microbial flocculant or biofocculant has emergedas a new research trend in flocculation technology. Generally, biofloccu-lation is a dynamic process resulting from synthesis of extracellularpolymer substances (EPS) by living cells (Salehizadeh et al., 2000). Upto now, bacteria, fungi and actinomyces have been identified asbioflocculant-producing microorganisms, in which these microorgan-isms are able to produce EPS, such as polysaccharides, functional pro-teins and glycoprotein, which act as bioflocculant (Gao et al., 2006). Arecent report has underlined that bioflocculant from Pseudomonasstutzeri and Bacillus cereuswas effective in flocculating marine microal-gae, P. carterae (CCMP647) (Lee et al., 2009). In the study, microalgaecells were not damaged even after flocculation. However, the studyalso pointed out that adequate mixing was essential to provide suffi-cient contact between EPS and microalgae.

In order to investigate the energy consumed during the mixingprocess, an assessment was carried to further strengthen the possibil-ity of bioflocculant in commercial scale (Lee et al., 2010a). In the as-sessment, baffled hydraulic flocculator was incorporated into themicroalgae harvesting system and the overall energy consumed wasestimated to be 0.9 kWh/tonne of dry mass flocculated or equivalentto 9.8 MJ/tonne biodiesel (assuming microalgae lipid content is 33%)(Lee et al., 2010a; Moheimani and Borowitzka, 2006). In addition,the cost estimated for this process was 79% lower compared to theconventional flocculant. Apart from that, flocculated culture mediumcan be effectively reused without retarding microalgae growth, there-by significantly reducing the cost for water treatment and purification(Kim et al., 2011). Nevertheless, research is still required to study theeffectiveness of using low-cost carbon source (e.g. glycerol) to cultureEPS producing microorganism.

Another possible method to harvest microalgae is through immo-bilization biotechnology, in whichmicroalgae are embedded in an en-trapment matrix and continuously grow within the matrix. Oncemicroalgae have grown to stationary phase and mixing is terminated,the immobilized microalgae beads will settled immediately at thebottom of the culture medium. Consequently, the relatively largesize microalgae beads can be easily separated from water throughsimple filtration method (e.g. sieve) that does not require highamount of energy. A conceptual culture system of immobilizedmicro-algae is illustrated in Fig. 3. Up to now, various immobilization tech-niques have been developed, however, only alginate gel entrapmentmethod is feasible to immobilize microalgae (Moreno-Garrido,2008). Alginate is a natural polysaccharide polymer (gelling material)which is extracted from brown algae. The polymer is mixed withmicroalgae cells and consequently stabilized with divalent ions such

Fig. 3. Visionary culture system of immobilized microalgae.


as Ca2+ to form immobilized microalgae beads through a nozzle(Mallick, 2006). Some of the advantages of using alginate gel are therequirement of only mild condition during immobilization process,negligible toxicity and high transparency (Moreira et al., 2006;Moreno-Garrido, 2008).

Immobilized microalgae beads can be applied in diverse researchareas such as for high-value products synthesis (e.g. H2), organicpollutants removal, heavy metal removal and toxicity measurement(biosensor) (Ertu rul et al., 2008; Guedri and Durrieu, 2008; Mallick,2006; Moreno-Garrido, 2008; Pérez-Martínez et al., 2010; Ruiz-Marin and Mendoza-Espinosa, 2008; Ruiz-Marin et al., 2010). How-ever, study on growing immobilized microalgae for biodiesel andbioethanol production is still scattered, especially in term of energyefficiency and cost saving evaluation. It is strongly believed thatsuccessful incorporation of immobilization technology in microalgaebiofuels production will eventually contributes a significant break-through towards the industry. Nevertheless, a few issues need to beaddressed in immobilization of microalgae before the process can beupgraded to commercialization stage: (1) Stability of the immobilizedbeads, especially when sea water or wastewater is used as culturemedium. High concentration of antigelling cations (e.g. sodium andmagnesium) in sea water and serious microbial attack in wastewaterwill cause dissolution of beads leading to loss of microalgae cells(Mallick, 2006; Moreira et al., 2006). (2) Leakage of microalgae cellsfrom immobilized beads. This problem normally occurs mainly dueto insufficient divalent ions during hardening process. Thus, the con-tinuous growth of microalgae inside the beads may cause the beads torupture. (3) Mass transfer limitation. After immobilization, a biofilmwill be formed to act as a protection layer and to restrict movementof microalgae cells. Consequently, nutrients and carbon source (main-ly CO2) need to diffuse through the biofilm before reaching the micro-algae cells. Hence, immobilized microalgae always recorded lowergrowth rate as compared to free cells culture. One of the possibleways to solve this problem is through co-immobilization with plant-

growth promoting bacteria (PGPB). PGPB such as Azospirillum brasi-lense was found to enhance growth of C. vulgaris significantly insmall alginate beads resulting to high microalgae biomass production(Gonzalez and Bashan, 2000). More explorations and studies of PGPBtowards other microalgae strains are strongly encouraged to facilitatethe development of microalgae immobilization technology in thefuture.

6. Drying of microalgae biomass

Unlike terrestrial energy crops, extensive drying of microalgaebiomass is required for biofuels production as the presence ofwater will inhibit several downstream processes, such as lipid ex-traction and transesterification. Nevertheless, not much concernhas been raised on this issue as solar drying is assumed to be thebest method to dry wet microalgae paste after the harvesting pro-cess. However, serious attention should be given as solar drying isnot feasible in temperate countries due to limited sunlight at certaintime of the year. In this case, heat generated from fossil fuels is re-quired to dry microalgae biomass continuously to ensure optimumbiomass production for each cycle of culture. However, a recentLCA study has highlighted that using natural gas as fuel for dryingmicroalgae biomass consumed nearly 69% of the overall energyinput and led to a negative energy balance in producing microalgaebiofuels (Sander and Murthy, 2010). In other words, heavy depen-dency on fossil fuels to dry microalgae biomass will seriously jeopar-dize commercial viability of microalgae biofuels and thus, newtechnologies or approaches (e.g. development of efficient dryers)are urgently needed to ensure the sustainability of microalgae biofu-el industry. Some other strategies such as to process microalgaebiofuels with partially or without drying through in-situ transesteri-fication and hydrothermal liquefaction will be discussed in detail inSections 8.3 and 9.


7. Microalgae lipid extraction

After harvesting and drying microalgae biomass, the subsequentstep is lipid extraction. Although the energy consumed in lipid extrac-tion from dried microalgae biomass contributed a relatively smallportion to the overall energy life cycle of microalgae biofuels (around5–10%) (Sander and Murthy, 2010; Stephenson et al., 2010), but thisprocess is still very important. Effective lipid extraction is requiredparticularly for microalgae with low lipid content as losing the lipidduring extraction process may bring a significant impact towardsthe production cost of microalgae biofuels (Ranjan et al., 2010).Different from terrestrial energy crops, lipid extraction from microal-gae biomass is relatively difficult due to the presence of thick cell wallthat prevents the release of intra-lipid. Hence, mechanical presswhich is effective to extract oil from terrestrial energy crops is gener-ally not applied to microalgae biomass. The following section depictstwo lipid extraction methods: (1) solvent extraction-suitable for drymicroalgae biomass and (2) supercritical fluid extraction-suitablefor wet-paste microalgae biomass.

Table 6Lipid extraction from various microalgae strains through chemical solvent.

Microalgae strain Chemical solvent Enhanced

Botryococcus braunii n-hexane: chloroform:methanol=4:2.67:1.33 (v/v/v)

Freeze-dr

DBUa Freeze-drDBUa:octanol=1:1 (mol/mol) Freeze-drDBUa:ethanol=1:1 (mol/mol) Freeze-dr

Botryococcus sp. Chloroform:methanol=1:1 –

Chloroform:methanol=1:1 AutoclaveChloroform:methanol=1:1 Bead-beaChloroform:methanol=1:1 MicrowavChloroform:methanol=1:1 UltrasoniChloroform:methanol=1:1 Osmotic

Chlorella Methanol: EMIMb=1.2: 1 (w/w) –

Chlorella protothecoides n-hexane Bead-bean-hexane:ethanol=1:1 (v/v) Bead-bea

Chlorella pyrenoidosa Chloroform:methanol=2:1 MagneticEthanol Magneticn-hexane MagneticChloroform:methanol=2:1 UltrasoniEthanol Ultrasonin-hexane Ultrasoni

Chlorella vulgaris Chloroform:methanol=1:1 –


Chlorococcum sp. n-hexane –

n-hexane/isopropanol –

n-hexane SoxhletChricystis minor Chloroform:methanol=1:2 –

Hot isopropanol –

Duniella Methanol: EMIMb=1.2: 1 (w/w) –

Microcystis Dimethyl ether –

Scenedesmus Chloroform:methanol=1:1 –


Scenedesmus dimorphus n-hexane Bead-bean-hexane:ethanol=1:1 (v/v) Bead-bea

Scenedesmus sp. n-hexane Soxhletn-hexane UltrasoniChloroform:methanol=3:1 (v/v) –

Chloroform:methanol=3:1 (v/v) Ultrasoni

a DBU=1,8-diazabicyclo-[5.4.0]-undec-7-ene.b EMIM=1-ethyl-3-methyl imidazolium methyl sulfate.

7.1. Solvent extraction

Chemical solvent extraction is the most common method used toextract lipid from microalgae biomass. This is because chemical sol-vent has high selectivity and solubility towards lipid and therefore,even inter-lipid can be extracted out through diffusion across micro-algae cell wall (Ranjan et al., 2010). Nevertheless, the disadvantagesof using chemical solvent are mostly related to their high toxicity to-wards human and surrounding environment. Chemical solvents suchas n-hexane, methanol, ethanol and mixed methanol–chloroform(2:1 v/v) (Bligh and Dyer method) are effective to extract microalgaelipid, but the extraction efficiency is highly dependent on microalgaestrains as shown in Table 6. From the table, modified Bligh and Dyermethod are the most favorable method to extract microalgae lipidfrom various strains and relatively high extraction efficiency can beattained compared to other solvents.

Although n-hexane is widely used to extract oil from various seedcrops, however, base in Table 6, it is inefficient to extract microalgaelipid. This is because microalgae lipid contains high concentration

method Lipid extracted(% of dry microalgae biomass)

Reference

y 2.7 Samorì et al., 2010

y 0.6 Samorì et al., 2010y 0.7 Samorì et al., 2010y 0.6 Samorì et al., 2010

8 Lee et al., 2010b11 Lee et al., 2010b

ter 28 Lee et al., 2010be 28.5 Lee et al., 2010bcation 8.5 Lee et al., 2010bshock 10 Lee et al., 2010b

38 Young et al., 2010ter 25 Shen et al., 2009ter 18 Shen et al., 2009strirring 19.7 D'Oca et al., 2011strirring 10.5 D'Oca et al., 2011strirring 2.5 D'Oca et al., 2011cation 19.4 D'Oca et al., 2011cation 5 D'Oca et al., 2011cation 1.5 D'Oca et al., 2011

5 Lee et al., 2010b10 Lee et al., 2010b

ter 8 Lee et al., 2010be 10 Lee et al., 2010bcation 6 Lee et al., 2010bshock 8 Lee et al., 2010b

1.5 Halim et al., 20115 Halim et al., 20115.7 Halim et al., 201122.9–26 Mazzuca Sobczuk and

Chisti, 201025–29.7 Mazzuca Sobczuk and

Chisti, 20108.6 Young et al., 201040.1 Kanda and Li, 20111 Lee et al., 2010b4 Lee et al., 2010b

ter 8 Lee et al., 2010be 10 Lee et al., 2010bcation 6 Lee et al., 2010bshock 5.8 Lee et al., 2010bter 30 Shen et al., 2009ter 24 Shen et al., 2009

0.8 Ranjan et al., 2010cation 0.9 Ranjan et al., 2010

2 Ranjan et al., 2010cation 6 Ranjan et al., 2010


of unsaturated fatty acid while n-hexane is a nonpolar solvent; thusthe selectivity of lipid towards the solvent is greatly reduced (Ranjanet al., 2010). Apart from that, it is worth to mention that n-hexane,methanol and chloroform are highly toxic compounds that can causesafety andhealth hazards if proper precaution steps are not taken. In ad-dition, it is not sustainable to use n-hexane and methanol since bothsolvents are conventionally derived from non-renewable fossil fuels.On the other hand, ethanol emerged as a greener solvent since it haslow toxicity level and can be derived from renewable sources such assugar-based plant (e.g. sugar cane and sweet sorghum) and lignocellu-losic material (e.g. wood and corn stover). However, base in Table 6,ethanol always give low extraction efficiency, mainly because ethanolis an azeotrop mixture (with 5% of water) and the presence of watermay possibly reduce its extraction efficiency.

For extraction using chemical solvent, diffusion is always the ratelimiting factor in the overall mechanism. However, this factor becomesmore serious in microalgae as the cell wall further prohibits solventfrom diffusing into the inner cell for lipid extraction. Therefore, cellsdisruption method can be introduced to enhance solvent diffusion effi-ciency and consequently, to improve microalgae lipid recovery rate.There are several techniques to disrupt microalgae cell wall, such asautoclave, bead-beater, ultrasonication and microwave as osmoticshock. All these techniques are depicted in Table 6. Although highermicroalgae lipid yield can be achieved after cell disruption, however,care should be taken as additional energy is required.

Table 7 shows a typical example of net energy analysis in microal-gae extraction system induced with different cell disruption methodsbase on a previous published report (Lee et al., 2010b). From thetable, negative energy balance is observed for autoclave and ultraso-nication, mainly because these machines have high energy consump-tion and the quantity of lipid recovered is relatively low compared tobead-beater and microwave. On the other hand, bead-beater attainedthe most promising result with the highest positive energy value,followed by microwave and non-disruptive method. The results sim-ply imply that introduction of cell disruption method in lipid extrac-tion does not always improve the system, instead it may lead tonegative net energy value. Nevertheless, it should be noted that theenergy analysis carried out in Table 7 is not an accurate comparisonsince proper optimization work are not performed and extractionefficiency is highly dependent on microalgae strains. Detail and com-prehensive LCA studies are therefore required to extend the findingsreported in Table 7, so that energy efficiency of different cell disrup-tion techniques towards the overall lipid extraction system can befurther justified. Apart from that, some of the cell disruptive methodssuch as ultrasonication and microwave posed several safety andhealth hazards and need to be addressed before up-scaling to com-mercial stage.

Table 7Energy efficiency in extracting lipid from Botryococcus sp. through various cells disrup-tive methods.

Cell disruptivetechnique

Lipid extracted(% of drymicroalgaebiomass)a

Total energyin microalgaebiodiesel(kJ)b

Additional energyconsumed due to celldisruptive process(kJ)

Netenergy(kJ)

Non-disruptive 8 1.51 0.00 1.51Autoclavec 11 1.89 4.28 −2.39Bead-beaterd 28 5.29 1.58 3.71Microwavee 28.5 5.39 2.09 3.30Ultrasonicationf 8.5 1.61 5.63 −4.02

a Data referred to Lee et al., 2010b.b A complete transesterfication reaction is assumed andmicroalgae biodiesel has en-

ergy content of 38 MJ/kg.c Power of 2 kW (capacity 35 L) is assumed to operate the machine (lab scale).d Power of 0.84 kW (capacity 80 g) is assumed to operate the machine (lab scale).e Power of 0.8 kW (capacity 28 L) is assumed to operate the machine (lab scale).f Power of 1.5 kW (capacity 20 L) is assumed to operate the machine (lab scale).

7.2. Supercritical fluid extraction

In the recent few years, research in extraction and reaction fieldhas entered a new dynamic era with the introduction of supercriticalfluid technology. The basic principal of this technology is achieving acertain phase (supercritical) that is beyond the critical point of a fluid,in which meniscus separating the liquid and vapor phases disappears,leaving only single homogeneous phase (Sawangkeaw et al., 2010).At supercritical phase, thermophysical properties such as density, vis-cosity, diffusivity and dielectric constant of a fluid will change drasti-cally depending on temperature and pressure. Consequently, thechanges of the thermophysical properties transform the fluid into asuper-solvent and thus, improve extraction and reaction efficiency.Several supercritical fluids that are currently being explored areethylene, CO2, ethane, methanol, ethanol, benzene, toluene andwater (Mendes et al., 2003; Sawangkeaw et al., 2010). Among these,supercritical-CO2 has received the most interest typically in extrac-tion of pharmaceutical and health related products from microalgae(Jaime et al., 2007; Kitada et al., 2009; Macías-Sánchez et al., 2008;Ota et al., 2009). In fact, supercritical-CO2 offers several advantagesin comparison with chemical solvent extraction: (1) non-toxic andprovide non-oxidizing environment to avoid degradation of extracts,(2) low critical temperature (around 31 °C) which prevent thermaldegradation of products, (3) high diffusivity and low surface tensionwhich allow penetration of pores smaller than those accessible bychemical solvents and (4) easy separation of CO2 at ambient temper-ature after extraction (Jaime et al., 2007; Mendes et al., 2003; Otaet al., 2009). However, the main disadvantages of supercritical-CO2

are associated with high cost of operation and safety related issues.Research studies on using supercritical-CO2 to extract microalgae

lipid for biodiesel production has been explored recently. In a study,lipid from wet-paste Chlorococcum sp. biomass was extracted usingsupercritical-CO2 and a lipid yield of 7.1% were attained at criticaltemperature of 60 °C, critical pressure of 30 MPa and extractiontime of 80 min (Halim et al., 2011). In addition, the lipid yieldattained from the wet-paste is even higher than dry biomass (5.8%),suggesting that energy consumed in drying process can be reducedthrough supercritical technology. Since supercritical-CO2 is a non-polar solvent, the presence of water in the system acts as a naturalpolar co-solvent and thus, facilitated the extraction of polar lipidsand improve total lipid yield extracted. Apart from that, soxhletextraction using hexane was found to be less efficient thansupercritical-CO2 extraction, achieving only 5.8% lipid yield after anextraction time of 330 min.

However, a contradicting result was observed when a comparativestudy between supercritical-CO2 and Bligh and Dye method (chemi-cal solvent extraction) were used to extract lipid from heterotrophiccultured microalgae C. cohnii (Couto et al., 2010). The lipid yieldattained from Bligh and Dye method was nearly double that of super-critical-CO2, indicating that microalgae strains and culture conditionsplays a significant role in determining the appropriate lipid extractionmethods. Although the energy consumed in operating supercritical-CO2 extraction is expected to be low due to the low critical tempera-ture of CO2, however, the energy required in separating pure CO2

from atmosphere and re-compressing the CO2 after each extractionshould not be ignored. Hence, a complete analysis is urgently re-quired to compare the feasibility of supercritical-CO2 and chemicalsolvent extraction in industrial scale, typically in term of energy effi-ciency and cost effectiveness.

8. Biodiesel production from microalgae

Lipid that is extracted frommicroalgae biomass is now ready to beconverted to biodiesel. The following sub-sections explain some ofthe possible methods to produce biofuels from microalgae lipid,such as transesterification using homogeneous and heterogeneous


catalyst and in-situ transesterification. Problems and suggestionsto improve each of the methods are included.

8.1. Homogeneous base and acid catalyst

The most commonway to produce biodiesel is through transester-ification, in which triglycerides (e.g. vegetable oils) are reacted withshort chain alcohol (e.g. methanol or ethanol) in the presence of cat-alyst as shown in Eq. (1) (Lam et al., 2010). Homogeneous base cata-lyst (e.g. KOH and NaOH) is usually used to accelerate the reaction.However, the presence of high free fatty acid (FFA) content in micro-algae lipid (more than 0.5% w/w) prevented the use of homogeneousbase catalyst for transesterification reaction (Ehimen et al., 2010; Zhuet al., 2008; Ziino et al., 1999). This is because FFA will react with basecatalyst to form soap leading to lower biodiesel yield and increase thedifficulty to separate biodiesel from glycerol (co-product). Alterna-tively, acid catalyst (e.g. sulfuric acid, H2SO4) will be a better optionas the catalyst is not sensitive towards FFA level in oil and thus, ester-ification (FFA is converted to alkyl ester) and transesterification canoccur simultaneously.

Base on currently available technologies, a two-step process in-volving combination of acid and base catalyst will be a better alterna-tive to enhance the feasibility of microalgae biodiesel conversionprocess. Microalgae lipid is initially subjected to acid pre-treatmentin order to reduce its FFA level before base catalyzed transesterifica-tion reaction takes place. This two-step process has been successfullyapplied in other biodiesel feedstock with high FFA content (e.g. wastecooking oil and jatropha oil) and can be easily up-scale for industrialapplications base on current established technology. Nevertheless,the main disadvantage of this two-step process is the use of extrabase catalyst to neutralize the acid catalyst during transesterification;an added cost to biodiesel production.

CH2-O-C-R1

CH2-O-C-R3

O

O

O

Triglyceride

+ 3CH3OH

CH3O-C-R1

O

CH3O-C-R2

O

O

CH3O-C-R3 CH2-OH

CH2-OH

CH-OH+

Methanol Methyl Ester Glycerol

CH-O-C-R2

ð1Þ

Up to now, optimization works on microalgae biodiesel conver-sion process using homogeneous base or acid catalyst is still verylimited in the literature. Most of the studies focus on up-stream pro-cesses, such as optimizing the phototrophic or heterotrophic cultureconditions, efficient photobioreactor design and effectiveness ofusing wastewater and CO2 from flue gases to culture microalgae. Sub-sequently, lipid composition is analyzed after a simple transesterifica-tion reaction using homogeneous base catalyst to quantify thesaturated and unsaturated fatty acids in microalgae biodiesel. Howev-er, reaction temperature, amount of catalyst used, alcohol to lipidmolar ratio and reaction time are not studied extensively, resultingto a void in knowledge on microalgae biodiesel conversion efficiency.A preliminary result has reported that microalgae lipid can be con-verted to biodiesel using similar reaction condition as other oilfeedstock (Plata et al., 2010); allowing the possibility to incorporatemicroalgae lipid to existing biodiesel plant without major modi-fication. From the study, more than 90% of microalgae biodiesel(C. vulgaris) yield can be attained at reaction temperature of 43 °C,methanol to oil molar ratio of 14, 0.42 wt.% of NaOH and reactiontime of 90 min. More research are deemed required, especially on

the effect of FFA in microalgae lipid towards the transesterificationreaction rate, potential and efficiency of other catalysts, microalgaebiodiesel properties and engine combustion performance.

8.2. Heterogeneous catalyst

Heterogeneous catalyst (base or acid) has also been exploredextensively for transesterification reaction to produce biodiesel. Un-like homogeneous catalyst, heterogeneous catalyst can be recycled,regenerated and reused for subsequently transesterification reactioncycles, thus enhancing cost effectiveness in biodiesel production.Furthermore, the catalyst can be easily separated out at the end of re-action through filtration and therefore, minimizing product contami-nation and number of water washing cycle (purification). To date,study on the application of heterogeneous catalyst in microalgae bio-diesel production is still limited, mainly because it is a relatively newfeedstock and not commercially available in the market. Commonheterogeneous base catalyst, CaO supported with Al2O3 was recentlytested for the very first time in transesterification of Nannochloropsisoculata microalgae lipid (Umdu et al., 2009). The highest microalgaeyield attained was 97.5% at reaction temperature of 50 °C, methanolto lipid molar ratio of 30:1, catalyst loading of 2 wt.% (referring toweight of lipid) and reaction time of 4 h. Long reaction time is alwaysrequired for heterogeneous catalyst since three immiscible phases(lipid–alcohol–catalyst) at the initial stage of reaction greatly in-creases mass transfer limitation within the system. In addition, itshould be noted that transesterification of microalgae lipid withpure CaO resulted to insignificant biodiesel yield whereas CaO sup-ported with Al2O3 (ratio 8:1 w/w) achieved the best result due tothe increasing number of basic density and basic strength of the cat-alyst. Other heterogeneous catalyst such as Mg–Zr and hierarchicalzeolites have also been investigated in transesterification of micro-algae lipid, however, unsatisfactory biodiesel yield was attained(less than 30%) (Carrero et al., 2010; Li et al., 2011b). Therefore,more breakthrough findings are urgently required to address the fea-sibility of heterogeneous catalyst in microalgae biodiesel industry.

8.3. In-situ transesterification

Conventional method to produce biodiesel mainly consists of twoseparate steps: extraction and followed by transesterification. In con-trast, in-situ transesterification simplify the process by allowing ex-traction and transesterification to occur in one single step, in whichoil/lipid-bearing biomass is directly contacted with chemical solventin the presence of catalyst. Chemical solvent plays two significantroles in this process: (1) as an solvent to extract oil/lipid out frombiomass and (2) as a reactant in transesterification reaction. In-situtransesterification offers several advantages over conventional bio-diesel production method such as minimize solvent separation step,reduce processing time and consequently, cut down the overall bio-diesel production cost (Shuit et al., 2010).

Furthermore, it is foreseen that in-situ transesterification canbring a greater advantage to produce biodiesel from microalgae bio-mass since mechanical extraction (e.g. extrusion and expeller) is inef-fective to extract lipid from microalgae biomass. In a typical study ofin-situ transesterification from dried Chlorella biomass, about 90% ofbiodiesel yield was attained at reaction temperature of 60 °C, metha-nol to lipid molar ratio of 315:1, H2SO4 concentration of 0.04 mol andreaction time of 4 h (Ehimen et al., 2010). Apart from that, it wasfound that methanol consumption can be reduced by introducingco-solvent during in-situ transesterification. Heterotrophic culturedChlorella pyrenoidosa with oil content of 56.2% was subjected to in-situ transesterification and 95% of biodiesel yield was attained withhexane as co-solvent (hexane to lipid molar ratio of 76:1), methanolto lipid molar ratio of 165:1, reaction temperature of 90 °C, 0.5 Mof H2SO4 and reaction time of 2 h (Miao et al., 2011). Other co-


solvents such as toluene, chloroform and dichloromethane were alsosuggested to be used during in-situ transesterification reaction (Xuand Mi, 2011).

Nevertheless, using wet-paste microalgae biomass has an adverseeffect in in-situ transesterification (Ehimen et al., 2010). In otherword, the wet microalgae biomass must be initially dried in orderto ensure efficient and optimum performance during in-situ transes-terification. A recent report has shown that microalgae biomasswith water content of more than 31.7% will completely inhibit thein-situ transesterification leading to negligible biodiesel conversion(Ehimen et al., 2010). A possible explanation to this phenomenon isthe occurrence of undesirable hydrolysis reaction during transesteri-fication. With the presence of water, triglyceride can be easily hydro-lyzed to diglyceride and FFA as shown in Eq. (2). Hence, instead ofextraction and transesterification, esterification of FFA also occursduring the reaction. Apart from that, it should be noted that metha-nol/ethanol are miscible in water. Therefore, during solvent recoveryprocess, water must be completely removed from the recoveredmethanol/ethanol before being used for subsequent in-situ transes-terification. Otherwise, the subsequent reaction cannot proceed atoptimum conditions due to the presence of previous solubilizedwater in methanol/ethanol as well as additional water in the freshmicroalgae biomass feedstock. Therefore, extensive biomass dryingis suggested prior to in-situ transesterification reaction to avoid theoccurrence of any side reaction and to simplify the subsequent sepa-ration processes.

CH2-O-C-R1

CH2-O-C-R3

O

O

O

+ H2O

CH2-O-C-R3

O

O

CH3-OH

+ HO-C-R1

O

Triglyceride Water Diglyceride Fatty acid

CH-O-C-R2 CH-O-C-R2

ð2Þ

9. Bio-oil production from microalgae

Since microalgae slurry contains high water content after harvest-ing, therefore, extensive drying is essential before the biomass is sub-jected to extraction and transesterification stage. Otherwise, thewater content will directly impede transesterification efficiency andcaused incomplete biodiesel conversion as discussed in Section 8.3.As aforementioned in Section 6, drying of wet microalgae biomassconsumed exceptional huge amount of energy typically in temperatecountries where sunlight is not available throughout the year.Furthermore, external heat which is usually generated from non-renewable sources (e.g. natural gas and coal) makes the drying pro-cess unsustainable for long term practice. In this regard, hydrother-mal liquefaction could be an alternative way to produce bio-oil frommicroalgae through aqueous-conversion method, in which freshlyharvested wet microalgae biomass are directly processed withoutdrying. Microalgae are expected to be an excellent biomass feedstockfor this technology because their small size will enhance rapid ther-mal transfer up to the required processing temperature (Heilmannet al., 2010). During hydrothermal liquefaction, water is heated tosub-critical condition (200 to 350 °C) under pressurized conditionin order to reduce its dielectric constant. The dielectric constant caneven drop to similar value as ethanol and thus, able to solubiliseless polar compounds (Duan and Savage, 2011; Kumar et al., 2011).In other words, water at sub-critical condition can serve as an effec-tive solvent but is significantly less corrosive than other chemical

solvents. Through hydrothermal liquefaction, satisfactory bio-oilyield and other gaseous products (e.g. H2) are attained from agricul-tural biomass and industrial waste such as sawdust (Karagöz et al.,2006), cattle manure (Yin et al., 2010), macroalgae (Zhou et al.,2010), grassland perennials (Zhang et al., 2009), wood (Zhong andWei, 2004) and secondary paper-mill sludge (Zhang et al., 2011).

Recently, several studies have investigated the potential of usinghydrothermal liquefaction technology to convert wet microalgae bio-mass to bio-oil and bio-char (Biller et al., 2011; Brown et al., 2010;Duan and Savage, 2011; Heilmann et al., 2010; Ross et al., 2010).43 wt.% of bio-oil was successfully recovered from Nannochloropsissp. (initial water content of 79 wt.%) through hydrothermal liquefac-tion at 350 °C for 60 min and the bio-oil obtained has a heating valueof 39 MJ/kg (Brown et al., 2010). However, the recovered bio-oil has arelatively higher composition of nitrogen and oxygen compared topetroleum crude oil, in which deoxygenation and denitrogenation isrequired to upgrade the bio-oil. More importantly, the process gavea positive energy of 45.3 kJ (assuming water enthalpies at 25 °C and350 °C is 82 and 1672 kJ/kg, respectively and the reactor is well insu-lated without any heat lost) indicating that hydrothermal liquefactionis a viable technology to convert wet microalgae biomass to bio-oilwithout requiring any drying process. On the other hand, the pres-ence of heterogeneous catalyst such as Co/Mo/Al2O3, Pt/Al2O3 andNi/Al2O3 during hydrothermal liquefaction are able to reduce oxygencomposition in bio-oil and increase its higher heating value (HHV) by10% (Biller et al., 2011). However, stability of the catalyst in term ofleaching, deactivation and regeneration under hydrothermal condi-tion is yet to be explored to justify the reliability of the catalysts forlong term usage. Apart from that, there are several issues that needto be addressed in hydrothermal liquefaction such as: (1) chemicalsolvent such as dichloromethane (DCM) is required to extract bio-oil from aqueous phase or thermal treated bio-char in which signifi-cantly reduce the process viability in industrial scale, (2) separationof the chemical solvent from aqueous phase and bio-oil is necessaryand possibly increase the overall energy input in the system, (3) theaqueous phase may contains high concentration of organic matterthat requires wastewater treatment before it can be discharged intowater sources and (4) there are more than 1000 different compo-nents in the bio-oil produced and therefore more research are re-quired to completely utilized all these components effectively.

10. Bioethanol production from microalgae

Up to now, only a handful of research information and data areavailable in the literature on the production of bioethanol frommicroalgae due to several reasons; (1) a lot of attention has beendiverted to biodiesel production frommicroalgae since certain strainsare capable to accumulate large quantity of lipid naturally inside theircells, (2) through nitrogen-deficient cultivation method (to save en-ergy and cost), lipid content inside the microalgae cells is boostedup significantly by blocking carbohydrate synthesis pathway, whilecarbohydrate is the main substrate to produce bioethanol and (3) bio-diesel has a higher calorific value than bioethanol, 37.3 MJ/kg and26.7 MJ/kg, respectively. Nonetheless, microalgae are found to be asuperior feedstock to produce bioethanol in comparison with otherfirst and second generation bioethanol feedstock. First generationbioethanol is derived from food feedstock such as sugar cane andsugar beet, in which over exploitation of this feedstock creates the“food versus fuel” issues and raised several environmental problemsincluding deforestation and ineffective land utilization. Second gener-ation bioethanol is produced from lignocellulosic biomass such aswood, rice straw and corn stover. Initially, these lignocellulosic bio-mass must be subjected to pre-treatment to break down the complexstructure of lignin and to decrease the fraction of crystalline celluloseby converting to amorphous cellulose (Cardona and Sánchez, 2007).However, most of the pre-treatment methods such as steam

Table 9Net GHG emissions of biodiesel production frommicroalgae, oil bearing crops and con-ventional diesel.

Biodiesel feedstock Net GHG emission(kg CO2-eq

a/MJ biodieselproduced)

Reference

MicroalgaeChlorella vulgaris 0.30b Stephenson et al.,

2010Chlorella vulgaris 0.04c Stephenson et al.,

2010Nannochloropsis salina −0.075d Batan et al., 2010Nannochloropsis sp. 0.32e Khoo et al., 2011Not specified 0.06f Clarens et al., 2010

Oil bearing cropCanola −0.05 Clarens et al., 2010Corn −0.08 Clarens et al., 2010Soybean −0.072 Batan et al., 2010Switchgrass −0.08 Clarens et al., 2010


explosion and alkali or acid pre-treatment are energy intensive andbring negative impact towards the environment.

In contrast, microalgae cells are buoyant and therefore, do not re-quire lignin and hemicelluloses for structural support (John et al.,2011). Hence, it is expected that the overall bioethanol productionprocess can be simplified due to the non-requirement of chemicaland enzymatic pre-treatment step. Nevertheless, it should be notedthat high concentration of carbohydrates are actually entrappedwithin the microalgae cell wall, in which an economical physicalpre-treatment process such as extrusion and mechanical shear isstill required to break down the cell wall so that the carbohydratescan be released and converted to fermentable sugars for bioethanolproduction (John et al., 2011).

On the other hand, simultaneous biodiesel and bioethanol produc-tion from microalgae is also possible, in which microalgae lipid isextracted prior to fermentation process. This concept has been prov-en viable in a recent study in which lipid from Chlorococum sp. wasextracted with supercritical CO2 at 60 °C and subsequently subjectedto fermentation by the yeast Saccharomyces bayanus (Harun et al.,2010a). From the report, microalgae biomass with pre-extractedlipid gave 60% higher ethanol concentration for all samples than thedried microalgae biomass without lipid extraction. This is becausesupercritical CO2 can act as a superior pre-treatment method tobreakdown microalgae cell wall causing the simultaneous release oflipid and carbohydrates embedded within the cell wall. Maximumbioethanol yield of 3.83 g/L was achieved from 10 g/L of lipid-extracted microalgae residue. In other words, lipid extraction frommicroalgae biomass for biodiesel production and pre-treatment stepto release carbohydrates for bioethanol production can occurs injust one single step which greatly enhanced the viability of microal-gae biofuels production in commercial scale. Apart from supercriticalCO2, other lipid extraction methods such as ultrasonication, chemicalsolvent, microwave and bead-beater have not been studied in orderto get a comprehensive comparison between the methods. Table 8presents the bioethanol yield from microalgae through several pre-treatment methods. The bioethanol yields obtained are comparableto the yields from sugary and lignocellulosic feedstock, indicatingthat microalgae biomass is a feasible alternative substrate for com-mercial scale bioethanol production (Choi et al., 2010; Harun et al.,2010a).

11. CO2 bio-fixation using microalgae

Microalgae are unicellular photosynthetic microorganisms thatcan convert solar energy to chemical energy with efficiency of10–50 times greater than terrestrial plants (Khan et al., 2009; Liet al., 2008b; Rosenberg et al., 2011). During photosynthetic process,microalgae utilized CO2 from atmosphere as carbon source to growand reproduce. Microalgae cells contain approximately 50% carbon,in which 1.8 kg of CO2 are fixed by producing 1 kg of microalgaebiomass (Chisti, 2007). Hence, this method is thought to be more en-vironmental friendly and technologically feasible to bio-mitigate CO2

Table 8Bioethanol production from different microalgae strains through various pre-treatment methods.

Feedstock Pre-treatment Ethanol yield(g ethanol/g substrate)

Reference

Chlorococum sp. SupercriticalCO2

0.38 Harun et al., 2010a

Chlorococumhumicolo

Acid 0.52 Harun and Danquah,2011

Chlorococuminfusionum

Alkaline 0.26 Harun et al., 2010b

Chlamydomonasreinhardtii

Enzymatic 0.24 Choi et al., 2010

compared to physicochemical adsorbents or direct injection into deepocean. However, the low concentration of CO2 in the air/atmosphere(0.04%) with poor mass transfer rate in water have resulted to theuse of expensive air pump to deliver CO2 efficiently to microalgaerather than relying on natural diffusion from atmosphere (McGinnet al., 2011). On the other hand, flue gases from industry usually con-tain more than 15% (v/v) of CO2 (Kumar et al., 2010) and therefore,could be a prospective source of carbon for microalgae. This is awin–win strategy in which air pollution from industry can be con-trolled through microalgae cultivation while the microalgae biomasscan be used to produce biofuels.

Currently, extensive research has been focused to identify suitablemicroalgae strains that can grow under high concentration of CO2

while producing lipid for subsequent biodiesel production. The de-sired microalgae strains should have the following characteristics:(1) high growth rate and biomass productivity, (2) high toleranceto trace amount of acidic components from flue gases such as NOx

and SOx and (3) able to sustain their growth even under extreme cul-ture conditions (e.g. high temperature of water due to direct intro-duction of flue gases) (Brennan and Owende, 2010). A few recentstudies have reported that Chlorella sp., Scenedesmus sp., and Botryo-coccus braunii are among the microalgae strains that have shownpromising result to bio-mitigate CO2 emission with typical CO2 con-sumption rate of 200–1300 mg/L/day (Chiu et al., 2008; Rosenberget al., 2011; Sydney et al., 2010; Yoo et al., 2010; Zhao et al., 2011).Apart from that, a pilot scale system was successfully developed toculture microalgae using industrial flue gases and Scenedesmusobliquus was able to tolerate high concentration of CO2 up to 12%(v/v) with optimal removal efficiency of 67% (Li et al., 2011a). Inaddition, supplying high concentration of CO2 to microalgae could en-hance the accumulation of polyunsaturated fatty acid in the microal-gae cells (Tang et al., 2011). This is an encouraging observation as

Fossil fuelConventional diesel 0.017 Batan et al., 2010

a CO2-eq is referred as CO2 equivalent.b Air-lift tubular bioreactor was used to culture microalgae. Flue gas with a content

of CO2 of 12.5 vol.% were supplied to microalgae as carbon source. Biodiesel with calo-rific value of 40 MJ/kg is assumed.

c Raceway pond was used to culture microalgae. Flue gas with a content of CO2 of12.5 vol.% were supplied to microalgae as carbon source. Biodiesel with calorificvalue of 40 MJ/kg is assumed.

d Closed-photobioreactor was used to culture microalgae. CO2 enriched air (2% CO2)were supplied to microalgae as carbon source.

e Integrated closed-photobioreactor with raceway pond was used to culture micro-algae. Compressed air enriched with 2% CO2 were supplied to microalgae as carbonsource.

f Raceway pond was used to culture microalgae. CO2 (not specified concentration)was supplied to microalgae as carbon source. GHG emission for extraction and transes-terification of microalgae lipid is not included in the calculation.


higher content of polyunsaturated acid tends to reduce the pour pointof biodiesel produced andmaking it feasible to be used in cold climatecountries.

Although there are a lot of benefits to culture microalgae for CO2

bio-mitigation, however, several recent LCA studies have shown oth-erwise (Clarens et al., 2010; Khoo et al., 2011; Stephenson et al.,2010). By referring to Table 9, most of the biodiesel derived frommicroalgae resulted to a net positive green house gas (GHG) emissioneven though flue gases or CO2 enriched air was used as carbon source.This is associated with the high power requirement in the cultivationstage; especially when air-lift photobioreactor was used to culturemicroalgae. Extensive amount of energy is consumed to operate theclosed photobioreactor for the following purposes: (1) continuouspumping of water to provide adequate mixing and (2) to enhancemass transfer rate of CO2 in water (Khoo et al., 2011; Stephensonet al., 2010). The energy required is normally generated from fossilfuels that emit a lot of GHG to the atmosphere. In contrast, microalgaecultured with raceway pond are less energy intensive than closedphotobioreactor with approximately 86% reduction in GHG emission.Another possible method to reduce GHG emission during microalgaecultivation is that microalgae farm should be located near powerplants for easy delivery of CO2 from the emitted flue gases. Alterna-tively, microalgae farming can be integrated with existing oil-bearing energy crops plantation as mentioned in Section 4. Excessbiomass from the energy crops plantation can be burned directly toprovide power for microalgae farming while the flue gases emittedwhich contains CO2 can be used as carbon source for microalgaegrowth. In short, the ability of microalgae to utilize CO2 effectivelyfrom flue gases is not the only factor that determine the global warm-ing potential (or GHG emission) of microalgae biofuels LCA, but otherfactors such as cultivation system and operating conditions are alsoequally important.

12. Conclusion

At the current stage, microalgae biomass is still not a viable choicefor commercial biofuels production due to the extensive energy inputcompared to current terrestrial energy crops. Base on several LCAstudies, the energy conversion efficiency ratio obtained for micro-algae is relatively lower than rapeseed, oil palm and jatropha, indicat-ing unsustainable biofuels production from microalgae. Severalenergy hotspots are identified in the overall microalgae processchain, including inorganic nitrogen source production, operation ofphotobioreactor and harvesting/dewatering of microalgae biomass.It is recommended that culturing microalgae for biofuels productionshould be coupled with wastewater treatment with the aim to mini-mize heavy dependency on inorganic nutrients source. Apart fromthat, incorporation of baffled system in open pond and closed-photobioreactor is suggested to enhance mixing intensity betweenmicroalgae, nutrient sources and CO2 while reducing the energyinput. In addition, effective harvesting and drying of microalgae bio-mass can be easily achieved through immobilization technology,however, extensive research is still required to strengthen this vision-ary strategy.

For the downstream processes, extraction of lipid frommicroalgaepresents a complicated task. Physical extraction method which issuitable to extract oil from oil bearing crops is not efficient in extract-ing lipid frommicroalgae since the lipid is embedded within a layer ofcell wall. Cell disruption method followed by chemical solvent extrac-tion is necessary to recover the lipid effectively. However, care shouldbe taken as some of the cell disruption methods require large quanti-ty of energy input that could lead to negative energy balance. In addi-tion, it should be noted that the choice of cell disruption methods,chemical solvents and extraction conditions are largely dependenton microalgae strains. In other words, there is no single methodthat can give optimum lipid extraction for all types of microalgae

strains. Several breakthrough technologies such as supercritical ex-traction/transesterification, in-situ transesterification, hydrothermalrecovery and transesterification assisted with ultrasonication or mi-crowave are yet to be discovered to enhance microalgae biodieselproduction.

Up to now, biodiesel production frommicroalgae still would ideal-ly be the main product. However, diversified biofuels production frommicroalgae is necessary to improve the overall energy balance. One ofthe successful examples is to use the microalgae biomass (after lipidextraction) for bioethanol production since high concentration of car-bohydrates still remain in the biomass. Other potential biofuels thatcan be derived from the microalgae biomass residue are, such asbio-oil from pyrolysis or hydrothermal process. This is a win–winstrategy in re-utilizing the waste to produce another source of energywhich greatly amplifies the sustainability of microalgae biofuels. Nev-ertheless, bioethanol and bio-oil production frommicroalgae is still atthe infancy stage and the real potential is yet to be completely discov-ered. Apart from that, an added advantage of culturing microalgae isits ability to bio-mitigate CO2 besides producing biofuels. Recent stud-ies have shown that it is technically feasible to utilize CO2 from fluegases as carbon source to culture microalgae. However, it is stillunclear if the presence of acidic gases (NOx and SOx) in the flue gaswill have any negative effect towards the growth rate of microalgae.In this regard, genetic engineering will have an important role inenhancing the overall life cycle of microalgae that are used as biofuelsfeedstock (Gong and Jiang, 2011; Greenwell et al., 2010; Huang et al.,2010; Radakovits et al., 2010; Rosenberg et al., 2008; Tabatabaei et al.,2011). Genetic modified microalgae that have the capability to growunder high concentration of CO2, able to withstand the presence ofcontaminants in flue gases or wastewaters and could produce highlipid content within their cells may be created. Advancement in thisresearch area is urgently required to bring a significant breakthroughin producing greener and sustainable microalgae biofuels.

For long-term sustainability and environmental benefits, all theprocessing stages of microalgae biofuels should be simplified withoutinvolvement of extensive energy input. In addition, the processesshould be easily adopted in the existing biofuels industry and canbe implemented immediately especially in third world countries.This is because culturing microalgae for biofuels production is notonly meant for profit making and benefiting the environment, butalso to help people from the bottom billions in terms of food andenergy security.

Acknowledgment

The authors would like to acknowledge the funding given by Uni-versiti Sains Malaysia (Research University Grant, PostgraduateResearch Grant Scheme (account number: 8044031) and USM Vice-Chancellor's Award) for this project.

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